Robust Double Emulsions for Multicolor Fluorescence-Activated Cell Sorting

Cell–cell interactions are essential for the proper functioning of multicellular organisms. For example, T cells interact with antigen-presenting cells (APCs) through specific T-cell receptor (TCR)–antigen interactions during an immune response. Fluorescence-activated droplet sorting (FADS) is a high-throughput technique for efficiently screening cellular interaction events. Unfortunately, current droplet sorting instruments have significant limitations, most notably related to analytical throughput and complex operation. In contrast, commercial fluorescence-activated cell sorters offer superior speed, sensitivity, and multiplexing capabilities, although their use as droplet sorters is poorly defined and underutilized. Herein, we present a universally applicable and simple-to-implement workflow for generating double emulsions and performing multicolor cell sorting using a commercial FACS instrument. This workflow achieves a double emulsion detection rate exceeding 90%, enabling multicellular encapsulation and high-throughput immune cell activation sorting for the first time. We anticipate that the presented droplet sorting strategy will benefit cell biology laboratories by providing access to an advanced microfluidic toolbox with minimal effort and cost investment.


Project background
The development and application of the analytical workflow described herein was a result of a collaboration between the microfluidics researchers at ETH Zurich (YD and AdM) and immunology researchers at the IRB Bellinzona (GZ, GA and RG).The IRB co-authors identified several limitations in existing droplet sorting and double emulsion (DE) sorting workflows, including the requirement for bespoke and expensive instrumentation, operational complexity, workflow inflexibility, and difficulties associated with multicellular sorting.Accordingly, the primary research objective was to establish a user-friendly workflow for DE sorting that addresses the aforementioned limitations.
The ETH researchers iteratively refined the analytical workflow based on continuous feedback from the IRB researchers, to realize a method that is both effective and accessible.The resulting workflow presented in the manuscript is now routinely used by cell biologists at the IRB.Indeed, all application-based data presented in the manuscript were independently conducted at the IRB.

Cells
Blood from healthy donors was obtained from the Swiss Blood Donation Center of Lugano and used in compliance with the Federal Office of Public Health (Authorization 2018-02166/CE 3428).Peripheral blood mononuclear cells (PBMCs) were isolated by Ficoll-Paque density gradient centrifugation (Sigma-Aldrich, Buchs, Switzerland).CD4 + and CD8 + primary human T cells were enriched from PBMCs with magnetic microbeads (Miltenyi Biotec, Adliswil, Switzerland) and cultured in RPMI-complete medium.Raji cells (ATCC, Manassas, USA) and Jurkat D1.1 cells (ATCC) were cultured in RPMI-complete medium.HEK293T cells (ATCC) were cultured in DMEM-complete medium.K562 cells (ATCC) were cultured in IMDM-complete medium.

Immortalization of primary human B cells with Epstein Barr Virus (EBV)
CD19 + B cells were enriched from PBMCs from healthy donors using magnetic microbeads (Miltenyi Biotec, Adliswil, Switzerland) and cultured in RPMI-complete medium.3x10 6 freshly isolated CD19 + B cells were inoculated with EBV (a kind gift from Antonio Lanzavecchia), cultured for 5 days, split and further expanded for 14 days.

Lentiviral transduction
Raji cells were transduced in 96-well plates (Corning, Corning, USA) (1.5 × 10 5 cells per well) in RPMI-complete medium.Concentrated lentiviral particles (pCDH-EF1α-MCS-(PGK-RFP)) were added, followed by spinoculation at 800 g for 30 minutes at 32°C.24 hours after transduction, cells were transferred to 48-well plates (Corning) and cultured in fresh RPMI-complete medium.After two days, RFP + cells were analyzed by flow cytometry and sorted using a FACS Aria III cell sorter (BD Biosciences, San Jose, USA).
Primary human T cells were activated with plate-bound anti-CD3 (5 μg/ml, clone TR66 (a kind gift of Antonio Lanzavecchia), anti-CD28 (1 μg/ml, clone CD28.2, BD Biosciences, San Jose, USA) in 96-well Nunc Maxisorb plates (Thermo Fisher Scientific, San Diego, USA) in the presence of IL-2 (400 U/ml, Thermo Fisher Scientific, San Diego, USA). 10 5 T cells were plated per well in RPMI-complete medium.After 24 hours of activation, concentrated lentivirus (pCDH-EF1α-MCS-(PGK-GFP)) was added and T cells were centrifuged at 800 g for 45 minutes at 32°C.24 hours later, cells were transferred to 48-well plates and cultured in IL-2-containing medium (50 U/ml).Five days post-transduction, cells were analyzed by flow cytometry and sorted using a FACS Aria III cell sorter.

Cell labeling
T cells were stained with anti-human CD2-PE (clone RPA-2.10;Biolegend, San Diego, USA) or anti-human CD45-APC (clone H130; Thermo Fisher Scientific, San Diego, USA) on ice for 30 minutes followed by extensive washing.Raji cells were labeled with either CellTrace Violet (CTV) or Cell Trace Far Red (CTFR), and K562 cells were labeled with CellTrace CFSE.All labels were sourced from Thermo Fisher Scientific, USA.Cells were resuspended at 10 6 /ml in PBS 1x and labeled with CTV, CTFR or CFSE (1:3,000 dilution) for 20 minutes at 37°C.Staining was quenched by adding five times the original staining volume of culture medium to the cells.After 5 minutes incubation, cells were centrifuged and resuspended in fresh medium.

Generation of NFAT-eGFP reporter
The sequence encoding for NFAT-eGFP was cloned from the retroviral plasmid pSIRV-NFAT-eGFP (Addgene plasmid #118031) into the lentiviral vector pCDH-EF1α-MCS-(PGK-NeoR) (System Bioscience, CD811A-1) that was modified as follows: to avoid constitutive expression of eGFP, the EF1α promoter was removed using NheI restriction enzyme digestion and the vector re-ligated.To insert the NFAT-eGFP sequence into the pCDH-MCS-(PGK-NeoR), donor and recipient plasmids were digested with BamHI and NotI enzymes.Digested DNA was separated on an agarose gel, excised, and purified using the NucleoSpin Extract II kit (Macherey Nagel, Oensingen, Switzerland).DNA ligation was carried out using T4 DNA Ligase (BioConcept AG, Allschwil, Switzerland) following manufacturer's instructions.All reactions were transformed into DH10B chemocompetent cells (Life Technologies Europe BV, Bleiswijk, Netherlands) and grown on LB+Agar plates containing 100 μg/mL Ampicillin (Roth AG, Arlesheim, Switzerland).Individual colonies were inoculated in liquid culture and plasmid minipreps were performed using NucleoSpin Plasmid kit (Macherey Nagel).Finally, insertion of the correct NFAT-GFP sequence was verified by Sanger sequencing (Microsynth AG, Balgach, Switzerland).

Microfluidic device fabrication
Both sets of microfluidic devices (for droplet generation and double emulsion conversion) were fabricated using polydimethylsiloxane (PDMS) following standard soft lithography protocols. 1The process involved structure design, mask fabrication, controlled exposure of the photoresist (photolithography), PDMS casting, bonding, and surface treatment, enabling the creation of desired channel structures in the PDMS devices.Briefly, a 2D projection of the microfluidic channel pattern was designed using CAD software (AutoCAD 2018, Autodesk, San Francisco, USA).The pattern was then printed onto a thin-film lithography mask by a third-party service (Micro Lithography Services, Chelmsford, UK).SU-8 3025 photoresist (MicroChem, Westborough, USA) was used to create features of different height (33 ± 1 μm for droplet generation chips and 43 ± 1 μm for DE conversion chips) on two silicon wafers by standard photolithography techniques. 2 Briefly, this involved exposing the spin-coated SU-8 molds to UV light through the lithography mask, followed by a development step to remove unexposed regions of photoresist.The SU-8 molds were then exposed to chlorotrimethylsilane (Sigma-Aldrich, Buchs, Switzerland) vapour for two hours to aid the detachment of PDMS replicas in a subsequent step.A 10:1 (wt:wt) mixture of PDMS base and curing agent (Sylgard 184, Dow Corning, Midland, USA) was thoroughly mixed and degassed and then poured over the SU-8 molds to form the top layers (~0.5 cm thick) of the microfluidic devices.After degassing to remove residual air bubbles, the molds and PDMS were cured in an oven at 70°C for two hours to ensure complete crosslinking of the polymer.Afterwards, the cured PDMS layer was peeled off from the molds and diced into individual devices.Inlet and outlet holes were created using a hole puncher (Syneo, Angleton, USA).The layers were then bonded to glass microscope slides (ground edge 45°, Thermo Scientific, Reinach, Switzerland) pre-coated with a 1 mm-thick layer of PDMS.Bonding was achieved by plasma treatment of PDMS surfaces using a plasma cleaner (Zepto Model 1, Diener Electronic, Ebhausen, Germany).This rendered the naturally hydrophobic PDMS temporarily hydrophilic, facilitating contact bonding process.To restore the hydrophobicity of the microchannel surfaces in droplet generation devices, they were placed in an oven at 70°C overnight.Conversely, the surfaces of channels within DE devices should remain hydrophilic.Accordingly, immediately after plasma bonding, a 1% (wt/wt) solution of polyvinyl alcohol (Mw 13,000-23,000, Sigma Aldrich, Buchs, Switzerland) in isopropyl alcohol (Sigma Aldrich) was injected into the microfluidic channels.The device was then left for 5 minutes to allow covalent linkage between the polyvinyl alcohol molecules and the PDMS, and formation of a permanent hydrophilic surface.Any excess solution was aspirated, and the device was placed on a hot plate at 120°C for 15 minutes to ensure a strong union between the PDMS layers.

Microfluidic co-encapsulation
Cells were counted using a FAST-READ 102 hemocytometer (Biosigma, Cona, Italy) and adjusted to the desired concentration in culture medium.Before loading into syringes, cells were filtered through a 40 µm mesh (Falcon cell filter, Fisher Scientific, Reinach, Switzerland) to remove cell aggregates. 1 ml BD Luer-Lok syringes (Fisher Scientific) were used to load the cell suspension (~ 0.3 ml per sample) and 5 ml BD Luer-Lok syringes (Fisher Scientific) were used to load the oil (Droplet Generation Oil for EvaGreen, Bio-Rad, Hercules, USA).To maximize cell viability, we opted not to use cell density gradient media to balance the cell suspension in our workflow. 3Instead, we arranged the syringes and pump in a vertical configuration for cell sample injection.Additionally, in the absence of a density gradient medium, cells naturally settle within the droplets, increasing the likelihood of cell interactions.It is worth noting that although cells gradually settle in the syringe over time, we observed no evidence of uneven cell encapsulation during a 90-minute period of operation.Masterflex Tygon tubing (ID 0.25 mm OD 0.76 mm, Fisher Scientific, Reinach, Switzerland) was used to connect the syringe needle (Gauge 27 TE tip, Distrelec, Nanikon, Switzerland) and microfluidic device.neMESYS syringe pumps (low pressure module, CETONI, Jena, Germany) were used to inject both oil and cell samples.Co-encapsulation was performed at a rate of 10 µl/min for the oil stream and 3 µl/min for each cell stream.Droplets were collected in another 1 ml BD syringe.During collection, a portion of the oil (in the lower layer) was aspirated off using a needle (Sterican 21G x 4 3/4", B. Braun, Melsungen, Germany) to provide sufficient space for the collection of all droplets.Other biocompatible fluorosurfactants, including RAN Biotech's 008-Fluorosurfactant, Dolomite's PicoSurf and Fluigent's dSurf, were tested for forming both droplets and DEs.All surfactants were used at 2% in Novec 7500 fluorinated oil.Tests (data not shown) indicated that all surfactants were compatible with our double emulsion workflow, with the only difference being small variations in the cell stream flowrate needed to maintain uniform droplet production.This is expected, since surfactants differ in their ability to reduce interfacial surface tension, which will directly impact droplet size. 4

Microfluidic DE conversion
After generation, droplets were stored for 30 minutes (with the outlet of the syringe facing upwards) before injection into the DE chip.This storage period allowed droplets to rise and accumulate at the syringe tip, ensuring that they entered the DE chip in a closely packed arrangement, thus minimizing the amount of accompanying oil.
The DE buffer, consisting of 2% (wt/wt) Poloxamer 188 (Sigma Aldrich, Buchs, Switzerland) and 1% (wt/wt) Tween 20 (Sigma Aldrich) in 1x PBS buffer (pH 7.4, Fisher Scientific, Reinach, Switzerland), was loaded into a 5 ml syringe.For DE conversion, we utilized the same pump and connection setup as for droplet generation.However, the pump was reoriented in the opposite (vertical) direction to match the adjusted position of the syringe.To prevent droplets from drying out, the tubing used for droplet injection was pre-coated with oil.This coating helps to prevent the transfer of oil away from the droplets, ensuring their stability and minimizing the likelihood of droplet drying and breakage.The droplet flowrate was varied between 1 and 4 μl/min and the droplet-to-buffer flowrate ratio was maintained at ratio of 1:4.An excessively slow buffer flowrate resulted in multiple droplets in a DE, while excessively high flowrates increased the likelihood of droplet splitting at the repacking orifice.

DE sorting into well plate
DEs in DE buffer were diluted 1:5 with PBS in a 12 × 75 mm round bottom FACS tube (BD Biosciences, San Jose, USA).DEs were gently resuspended by manual swirling of the tube.To maximize DE sorting efficiency, the droplet delay was adjusted from -1.5 to +1.5 delay units (droplet cycles) in increments of 0.25-0.5 delay units.For each delay unit, 1-2 droplets were sorted into different wells of a 96-well plate containing DE buffer.Subsequently, DE drops were imaged using an ImageXpress Micro 4 high-content microscope (Molecular Devices, San Jose, USA) with a 10x objective.One image in both brightfield and fluorescence channels was taken for each well using an exposure time of 200 ms.Finally, wells containing one or more DE drops were manually counted using a benchtop microscope to assess sorting efficiency.All FACS data were analyzed using FlowJo v10 software (BD Biosciences, Ashland, USA) for quantification and assessment of the sorting process.

Droplet imaging
To evaluate co-encapsulation efficiency, primary human T cells expressing GFP and CTV-labeled Raji cells were resuspended in RPMI-complete medium at different concentrations (1 × 10 6 /ml, 2 × 10 6 /ml, 4 × 10 6 /ml, 8 × 10 6 /ml and 15 × 10 6 /ml) and co-encapsulated (oil flowrate = 10 µl/min, cell flowrate = 3 µl/min).Encapsulation was monitored on an E800 upright microscope (Nikon Instruments, Shinagawa, Japan) with a 4x objective, brightfield illumination and MotionBlitz EoSens mini1 high-speed camera (Mikrotron, Unterschleissheim, Germany).To capture higher levels of detail, additional tests were conducted using a Nikon Eclipse Ti-E Inverted microscope (Nikon Instruments) equipped with a Phantom Miro M310 high-speed camera (Vision Research, Wayne, USA).Following co-encapsulation, imaging of W/O droplets was performed using a TE300 inverted microscope (Nikon Instruments, Shinagawa, Japan) with a 10x objective, bright-field illumination and acquired using a Hamamatsu C9100 EM-CCD camera (Hamamatsu Photonics, Hamamatsu City, Japan).Four representative brightfield images were taken for each condition and analyzed using ImageJ/Fiji (https://fiji.sc).To count the total number of droplets, images were first subjected to a "Gaussian Blur" smoothing filter and subsequently analyzed with the "find maxima" function.Droplets containing one or more cells were manually counted using the "multi-point function" in ImageJ/Fiji.

External osmolarity
To assess the effects of external osmolarity, GFP-expressing primary human T cells and CTV-labeled Raji cells were separately resuspended in RPMI-complete medium at a concentration of 4 × 10 6 /ml and co-encapsulated.Droplets were then double emulsified.Four sets of DE drops were incubated in 1x, 2x, 4x and 8x of PBS for 1 hour at 37°C.Before and after sorting, DE drops were analyzed with the Nikon TE300 microscope with 10x objective, Nikon Hg fluorescence lamp (Nikon Instruments, Shinagawa, Japan) and Hamamatsu C9100 EM-CCD camera.Images were acquired in the bright field and DAPI channel with an exposure time of 200 ms.

Chip design
The three chip designs used are provided below.The original CAD file with exact dimensions is provided separately.

Thin-shell DE advantages
The shear stress, τ, in the shell is inversely related to the thickness, h, while the flow velocity along the boundary and the dynamic viscosity of the liquid are represented by U and μ, respectively.In rotational flows, a deflection angle is created between the oil shell and water core, resulting in a Laplace pressure (ΔP) "shock". 7,8he surface tension of the interface and the radius are denoted by σ and R, respectively.Bottom panel adapted with permission from the Royal Society of Chemistry. 8

DE stability tests
Cell-laden DEs showed good stability during storage and exposure to mechanical disturbances.DEs stored for a period of 15 months exhibited negligible rupture.DE resilience to acute mechanical stress was further assessed by vortexing DEs at maximum speed for one minute using a Mixer Vortex Genie 2 in an Eppendorf tube.DEs remained intact after this disturbance test.Detailed results from these stability tests are presented in Figure S3.We evaluated DE stability to ionic strength variations.Specifically, we replaced the original DE buffer with fluids of variable osmotic pressure: distilled water (hypotonic), PBS and cell media (isotonic), and 10x PBS (hypertonic), as detailed in Figure S4.Data indicated excellent DE stability under both isotonic and hypertonic conditions.Interestingly, whilst internal droplets within DEs shrank when using 10x PBS, this did not compromise DE stability.Conversely, in distilled water, DEs were swelled but remained intact.However, it was noted that when droplets included 16% OptiPrep, a common additive used to control cell density, over half the DEs burst (Figure S8c).Given the obvious stability of DEs across various media, we propose substituting external buffers with the relevant cell culture medium for prolonged cell cultivation.Previous studies have identified three primary mechanisms for material exchange across the oil layer in DEs, namely diffusion, surfactant-based chemical extraction, and surfactant-assisted transport. 5Given its thin oil shell, our DE system is likely to enhance such material exchanges, potentially facilitating more extended cell culture periods.However, further research is required to conclusively determine these benefits.To evaluate the stability of DEs under the application of electric fields, we assessed DE-protected droplet merging events in a microfluidic channel.As shown in Figure S5, paired droplets (one containing PBS the other containing 10% blue ink) in a DE were passed through a microfluidic channel.No droplet merging events were observed as a function of voltage and frequency within the tested range (100-1000 V, 1-10 kHz).For comparison, we also assessed the merging of "normal" droplets under similar electric field conditions in a microfluidic channel (Figure S5).More in depth analyses of (microfluidic) droplet merging in electric fields can be found elsewhere. 9,10

Osmotic pressure manipulation
Osmotic pressure control is an effective method for altering DE size post-formation. 11A disparity in osmotic pressure between the internal and external aqueous phases will initiate water transport across the oil shell of a DE, leading to either DE shrinkage or swelling. 12,13Water transport mechanisms encompass direct diffusion through the oil layer and transport via "reverse micelles"; the latter being predominant structures formed by surfactants in the oil phase. 14,15We hypothesized that reducing the internal water core might boost the chances of two-cell interactions.Interestingly, a recent study by Zhuang et al. also suggested that DE shrinkage can be used to increase biomarker concentration in the core, thereby enhancing signal detection. 16To assess the feasibility of implementing DE contraction and its impact on FACS detection performance, we divided a batch of cell-laden DEs into four groups and replaced their buffers with solutions of 1x PBS, 2x PBS, 4x PBS, and 8x PBS, respectively.
As expected, significant shrinkage occurred.Compared to 1x PBS, DEs in 8x PBS exhibited a 30% reduction in their final diameter (Figure S9e and f).Although water migration across DEs can occur over several hours, 12 employing a thin oil layer significantly accelerates this process. 17Consequently, in the current experiments, migration was swift, taking only a few minutes.FACS analysis shows how varying osmotic pressures affect DE populations (Figure S9a-d).With a decrease in DE size from 1x to 8x PBS, we observed a corresponding drop in the scatter-gated DE population from 92.3% to 69.1%.This reduction is likely due to difficulties in distinguishing contracted DEs from empty oil droplets and other debris.In contrast to the findings of Zhuang et al., 16 where signal enhancement in fluorescent cellular secretions was observed with DE contraction, our study revealed no clear correlation between fluorescence detection rates and DE shrinkage.Specifically, CTV and GFP detection rates varied minimally across 1x to 8x PBS concentrations.This inconsistency implies that signals emanating from the cell surface or interior do not become enhanced by DE volume reduction.Accordingly, it is essential to consider the signal source when manipulating DE size, as benefits to signal detectability may not be universally applicable.

Figure S2 .
Figure S2.Molecular transport and hydrodynamic properties of DE drops.(a) Mechanisms for molecule transport across the DE oil shell include simple diffusion, surfactant-based chemical extraction, and surfactantassisted transportation. 5A thin shell accelerates mass transfer by shortening the transit distance and increasing the concentration gradient.Panel adapted with permission from the Royal Society of Chemistry. 5(b) Hydrodynamic models for DE drops in shear and rotating flows.Within shear flows, internal circulating flows are formed in the oil shell (yellow zone) and water core (white zone) of a DE drop, and caused by friction between the immiscible phases.6,7The shear stress, τ, in the shell is inversely related to the thickness, h, while the flow velocity along the boundary and the dynamic viscosity of the liquid are represented by U and μ, respectively.In rotational flows, a deflection angle is created between the oil shell and water core, resulting in a Laplace pressure (ΔP) "shock".7,8The surface tension of the interface and the radius are denoted by σ and R, respectively.Bottom panel adapted with permission from the Royal Society of Chemistry.8

Figure S3 .
Figure S3.Assessment of the durability and mechanical stability of cell-laden DEs.(a and b) DE storage stability over 15 months.Cells were fixed in 4% formalin prior to encapsulation within DEs, which were stored initially at 4°C for three months and then room temperature for twelve months.DE diameter distributions are shown for (a) freshly formed DEs (CV 3.66%) and (b) aged DEs after 15 months (CV 6.54%).Overlapping DEs and DEs at image edges were excluded from analysis in (b).(c) Mechanical resilience of DEs post-vortexing.DEs containing live cells were subjected to intense vortexing for one minute at a power setting of 10 (maximum speed) on a Mixer Vortex Genie 2. The DE diameter distribution post-vortexing exhibited a CV of 5.31%.Images in (b) and (c) were captured through the eyepiece of a microscope using a smartphone camera.Due to lens distortion, extracted CV values in phone images are likely to be overestimated.Scale bars in all images are 50 µm.

Figure S4 .
Figure S4.Incubation stability test.DE drops (contained cell media) are incubated in different osmotic concentrations.(a-c) Isotonic: original DE buffer, cell media and 1x PBS.(d) Hypotonic: DI water.(e) Hypertonic: 10x PBS.No breakage is observed in all test environments.The scale bars in all images are 50 μm.

Figure S5 . 8 . 14 9.
Figure S5.DE stability under alternating electric fields.(a) The DE shields contents from an external electric field.DEs consisting of paired droplets pass through a periodically squeezed channel.The schematic illustrates the experimental setup integrating salt electrodes and the DE flow path.Brightfield images present DEs under an applied electric field of 500 Vpp at 2 kHz, with no droplet merging being observed.The associated video is provided as Video S3.The map plot shows the tested voltage and frequency ranges for DE shell-protected droplet merging experiments.(b) Normal droplet merging under a similar electric field condition in a microfluidic channel.The sequence of images depicts the merging of normal droplets under an electric field of 500 Vpp at 2 kHz over 3.5 ms.The scale bars in all images are 50 μm.

Figure S9 .
Figure S9.The impact of DE external osmolarity.GFP-expressing T cells (4 million/ml) and CTV-stained Raji cells (4 million/ml) were co-encapsulated and double emulsified before being incubated in PBS buffers of varying concentration.Flow cytometry analysis was then conducted to evaluate the effects of different osmolarities: 1x PBS (a), 2x PBS (b), 4x PBS (c), and 8x PBS (d).The size of the resulting DEs was observed under a microscope (e), and trend plots of DE diameter and gated DE proportion relative to the PBS concentration are shown in (f).The scale bar in the image is 50 μm.